Characterization of Wolbachia pipientis Infections in Arthropods Native to Various Coastal
New Jersey Habitats
Joshua D. Nelson, Felicity Bennett, Lee Kerkhoff, PhD
Submitted to Marine Biological Labs to fulfill the requirements for the MBL Summer
Envisionship Program
Introduction
Wolbachia are an abundant and highly diverse group of endosymbiotic alphaProteobacteria, related to the Anaplasma, Erlichia, and Rickettsia genera, that was first described
as a parasite in the mosquito Culex pipientis by Hertiz and Wolbach in 1924. After its discovery,
Wolbachia went virtually unstudied until the early 1970s (1-3, 8, 9). Wolbachia was considered
somewhat rare until the 1990s, when molecular biological techniques allowed investigators to
discover that Wolbachia were widespread and very abundant in diverse hosts including
arthropods and nematode worms. Since then, Wolbachia bacteria have been studied extensively,
aided largely by further advances in molecular biology methods(1, 2, 5, 13, 18, 25). To date,
eight distinct Wolbachia “supergroups” have been characterized, designated by letters A through
H, based on 16S rRNA as well as functional gene sequence data (1-7). Wolbachia has become
known as the most widespread obligatory endosymbiont on the planet, and field sampling and
statistical analyses estimate that 20 to 60% of arthropod species are infected (10, 25).
Despite recent advances in knowledge about Wolbachia, relatively little is known about
the phylogenetic and evolutionary history of these bacteria, due to lack of DNA sequence data
and challenges in identifying, or “strain-typing” these bacteria (2, 5). Baldo et al (2006) recently
described multi-gene approach identifying Wolbachia strains using five genes that are highly
conserved and that is ubiquitous to the Wolbachia genus, as well as a gene marker for a
Wolbachia surface protein (2). This work has led to the formation of a Wolbachia strain-typing
database that continues to grow.
These amazing and diverse endosymbiotic bacteria are transmitted vertically through host
germ cells and have evolved endosymbioses ranging from mutualism to reproductive parasitism.
For example, Wolbachia have evolved mutualistic relationships with filarial nematode hosts,
which depend on Wolbachia for survival and likely protect nematode worms from host immune
systems (5, 18, 23, 25). In contrast, Wolbachia have evolved reproductive parasitism in
arthropod hosts, and elicit a variety of phenotypic effects (Figure 1) including feminization,
parthenogenesis, male-killing, and sperm-egg incompatibility (cytoplasmic infertility, also
referred to as “CI”) (8, 90).
Figure 1. A summary of Wolbachia-induced phenotypic effects in arthropods (from Werren et
al, 2008) (25).
Several studies have shown that the phylogeny of nematode-associated agree closely with
the phylogeny of the host species. Filarial nematodes are infected by only two Wolbachia
supergroups, C and D(5, 18, 23, 25). This suggests little horizontal movement of Wolbachia
among nematode species. In contrast, Wolbachia infections in arthropod hosts show little
agreement between host phylogeny and Wolbachia group phylogeny, suggesting extensive
horizontal transmission within and between arthropod hosts. Host phenotypic responses,
described above as feminization, parthenogenesis, male-killing, and sperm-egg incompatibility,
vary depending on the host arthropod species. In other words, a Wolbachia strain that infects
one insect taxa, causing CI, may cause male-killing in another insect taxa (1, 2, 5, 18, 23, 25).
Casiraghi et al demonstrated that although Wolbachia supergroups are restricted to either
arthropods or nematodes, one supergroup was found in both, supporting several theories on
horizontal movement of Wolbachia between highly divergent invertebrates (6). These results
agree with the results of Bordenstein and Rosengaus (2005), which described a novel supergroup
of Wolbachia in Isopterans that is distinct from previously described Wolbachia that infect
Isopterans, suggesting horizontal transmission rather than a single acquisition (5). Figure 2
summarizes the phylogeny of Wolbachia supergroups. While the mechanism of Wolbachia
horizontal transmission is not well characterized, Kyei-Poku et al. (2005) proposed one
mechanism, by which parasitic wasps that feed on other arthropods could acquire a Wolbachia
infection and spread it to other uninfected hosts (13).
Figure 2. Summary of Wolbachia supergroups (from Werren et al, 2008) (25).
The study of Wolbachia in the last fifteen years has revealed several unique traits that
previously have not been described in other endosymbiont microbes. Wolbachia has is an
exception to the rule of endosymbiosis, as it undergoes extensive recombination, both within and
between super-groups,(1, 2, 24, 25) resulting in exceptional levels of diversity among Wolbachia
strains. Baldo et al (2006) reported divergence of 6.5 to 9.2% among Wolbachia strains, and
have identified 37 unique strains.
Why study Wolbachia?
In addition to its abilities to extensively recombine between strains, horizontally transmit,
and influence host evolution, Wolbachia has become known as the causative agent in
pathogenicity of parasitic nematode worms(15). Several diseases, including Lymphatic
filariasis, river blindness, chronic inflammatory disease, and elephantitis, are associated with
nematode worm infections, and 200 million people worldwide are affected, representing an
immense global public health concern (15, 18, 19).
While the exact mechanism of Wolbachia’s role in filarial disease has not been
fully characterized, data from several studies suggests that Wolbachia is essential to the survival
of various parasitic nematode worms and that the immune response to filarial worm infections is
likely a response to the release of endo-toxin-like molecules released when worms die or
degenerate (15, 18, 21, 25). Antibiotic therapy used in conjunction with traditional anti-filarial
treatment has been shown to affect nematode embryonic development, stunt larval development,
kill adult worms. Treatment with anti-filarial drugs alone has also been shown to elicit
inflammatory response in treated patients, suggesting a Wolbachia-antigen that may be linked to
inflammatory disease. These discoveries have led to treatments that decrease host inflammatory
disease and ocular inflammation (15, 18, 21, 25).
As a result of several recent studies, Wolbachia has been increasingly studied as a target
for elimination and treatment of filarial nematode infections as well as treating inflammatory
disease in human and livestock hosts. In addition, Wolbachia are currently being studied as
agents for controlling pest arthropod populations and insect-borne diseases (15, 16, 18, 21, 25).
For example, the vertical transmission of Wolbachia could be used to insert “transgenes”, or
foreign genes, into Wolbachia to drive selected traits in an arthropod vector species (16).
Alternatively, infected arthropod species could be used to increase cytoplasmic incompatibility
in a population, thereby increasing mortality in pest arthropod species (4, 8, 17, 23-26).
However, while these mechanisms show promise, reliable manipulation of Wolbachia in this
way has yet to be achieved, and controlling filarial nematode populations using Wolbachia has
proven more successful.
In addition to being instrumental in controlling filarial nematode disease and controlling
arthropod populations, the potential for using Wolbachia strains models for understanding
evolution seems limitless. Wolbachia may be responsible for a variety of evolutionary
mechanisms including host speciation, reproductive isolation, lateral gene transfer from a
prokaryotic endosymbiont (Wolbachia) to a eukaryote host, and bacterial recombination (3, 4,
10, 25).
Materials and Methods
Insect collection
Insect samples were collected at random from five different sites using a previouslydescribed sweep-net technique (20, 22). Sampling was conducted for two hours each time and
was done both during the morning hours (between 8am and 11am) and during the late afternoon
(between 2pm and 5pm).
Sampling site information and site descriptions are summarized below:
Site:
Point Pleasant, NJ
Backyard
Site Description
Suburban neighborhood. Sunny area with lawn grass and long
flowering annual and perennial plants. There is very little shade
and is very dry.
Wall Township, NJ
Backyard
Damp with maple and oak trees and thick bush. Very little
grass grows in the shady area but grass is tall in the sunny areas
and large patches of ivy cover most of the uncovered ground.
Spring Lake Hts., NJ
Backyard
Suburban neighborhood. Sunny area with lawn grass and long
flowering annual and perennial plants. There is very little shade
and is very dry.
Wreck Pond, Wall, NJ
Riparian area that is part of a watershed that feeds into the
Atlantic Ocean. The area is covered with dense low shrubs,
pine, maple, and cedar trees, grasses, and sedges.
55-acre State Park that is part of the Manasquan River Estuary
that feeds into the Atlantic Ocean. The park size is comoprised
of saltmarsh areas and areas of dense low shrubs, pine, maple,
and cedar trees, grasses, and sedges.
Fisherman’s Cove,
Manasquan River,
Manasquan, NJ
Allaire State Park:
3200 acre park comprised of dense forest composed of oak and
maple species. Also includes river floodplain areas and riparian
areas. These samples were collected from a wooded area near a
small lake near a the main nature center in the park. The ground
is sandy and covered in a layer of decaying leaves and other
materials.
Samples were netted and placed immediately in 95% ethanol then placed in a -20 degree
Celcius freezer within twenty minutes of the end of the sampling session. Samples were stored
at -20 degrees Celcius in 95% ethanol until sample sorting and washing.
Insect samples were sorted and washed using the MBL/Discover the Microbes Within
insect identification lab protocol (7). Samples were sorted into various morphospecies (Table 1)
per the MBL/Discover the Microbes Within insect identification lab protocol and were processed
rapidly to maintain DNA quality. Redundancies and cross-referenced species were noted, where
applicable and species identification was made, when possible. Furthermore, voucher species
were preserved frozen in 95% ethanol for future species identification. Sorted samples were
placed back in the -20 degree Celcius freezer until DNA extraction was performed.
DNA Extraction
DNA extractions were performed from a minimum of one and a maximum of eleven
samples from each representative morphospecies per the MBL/Discover the Microbes Within
DNA Extraction protocol and the Qiagen DNeasy tissue DNA extraction protocol (7). Samples
were chosen at random for DNA extraction. If the insect measured larger than 2-5mm, a 2-3mm
section was removed from the insect abdomen for DNA extraction. Otherwise, the entire insect
was used for DNA extraction. Samples were completely ground/pestled, per the Qiagen DNeasy
protocol and were processed exactly according to the protocol to ensure DNA quality. Genomic
DNA samples were run on 1% agarose gels for the first two sets of DNA extractions to
qualitatively test for DNA presence and quality.
PCR Amplification and Gel Electrophoresis
The MBL/Discover The Microbes Within PCR protocol was used to amplify a Wolbachiaspecific 438 base-pair fragment (WSPEC) of the prokaryotic 16SrRNA gene and a 658 base-pair
fragment of the eukaryotic cytochrome oxidase (CO) gene (7).
PCR was carried out in 25µl volumes using the GE-Healthcare PCR bead kit and the
MBL/Discover The Microbes Within PCR protocol. Three controls were used for each set of
samples amplified. The controls are summarized below:
(+) Nasonia – a known Wolbachia-positive sample
(-) Nasonia – a known Wolbachia-negative sample
No-DNA – a sterile-milliQ water negative control.
PCR was conducted using the following primers:
WSPEC Forward: (5’-CATACCTATTCGAAGGGATAG-3’)
WSPEC Reverse: (5’-AGCTTCGAGTGAAACCAATTC-3’)
CO Forward (are LCO1490): (5’-GGTCAACAAATCATAAAGATATTGG-3’)
CO Reverse (HCO2198): (5’-TAAACTTCAGGGTGACCAAAAAATCA-3’)
PCR cycle information is summarized below:
1 cycle
1 min @ 94 C
38 Cycles
1 min @ 94 C
1.5 min @ 45 C
1 min @ 72 C
1 cycle
5 min. @72 C
Each PCR product was run on 1% agarose using 1X TAE buffer. 10µl of each PCR product was
electrophoresed for 48 minutes at 75 volts in a chilled gel-boat. PCR product size was
determined using a 1kb ladder. Gels were stained with ethidium bromide and then visualized in
UV light. PCR products were frozen immediately after PCR cycling was completed to ensure
DNA quality. Positive Wolbachia infections were identified and recorded in the results section.
Results
A total of 96 unique insect samples were PCR tested for both the eukaryote CO gene
fragment and the prokaryote Wolbachia-specific 16SrRNA fragment. 29% of all samples tested
were positive for the Wolbachia-specific fragment, in close agreement with previous estimates
(10, 23-25) PCR results are summarized below:
Table 1: Morphospecies list and PCR test results. Positive results indicate a positive Wolbachiaspecific 16SrRNA fragment.
Morphospecies
FB2010-17
Order
Hymenoptera
FB2010-21-1
Hymenoptera
FB2010-21-2
Hymenoptera
FB2010-22-1
Hymenoptera
FB2010-22-1 dup pcr
Hymenoptera
FB2010-22-2
Hymenoptera
FB2010-22-2 dup pcr
Hymenoptera
Date
collected
7/19/10
7/20/10
7/20/10
7/20/10
7/20/10
7/20/10
7/20/10
FB2010-26-1
Diptera
7/22/10
FB2010-26-2
Diptera
7/22/10
FB2010-28
Hemiptera
7/22/10
FB2010-28 dup pcr
Hemiptera
7/22/10
FB2010-31
Orthoptera
7/22/10
FB2010-33
Diptera
7/22/10
FB2010-34-1
Hemiptera
7/22/10
FB2010-34-2
hemiptera
7/22/10
FB2010-36
Hemiptera
7/22/10
FB2010-39
Hymenoptera
7/22/10
FB2010-39 dup pcr
Hymenoptera
7/22/10
FB2010-40-1
Hemiptera
7/22/10
FB2010-40-2
Hemiptera
7/22/10
FB2010-41-1
Diptera
7/22/10
FB2010-41-2
Diptera
7/22/10
FB2010-42
Diptera
7/22/10
FB2010-46-1
Hymenoptera
7/22/10
FB2010-46-1 dup pcr
Hymenoptera
7/22/10
FB2010-46-2
Hymenoptera
7/22/10
FB2010-47
FB2010-47
Hymenoptera
Hymenoptera
7/22/10
7/22/10
Location
Pt. Pleasant
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
PCR test
+
+
+
+
Species/Description
JDN2010-27-1
Hymenoptera
7/20/10
JDN2010-27-2
Hymenoptera
7/20/10
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Point
Pleasant
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Allaire State
Park
Spring Lake
Hts
Spring Lake
Hts
JDN2010-29-1
Coleoptera
7/20/10
Wreck Pond
Tiny brown beetle
JDN2010-29-2
Coleoptera
7/20/10
Wreck Pond
Tiny brown beetle
JDN2010-29-3
Coleoptera
7/20/10
Wreck Pond
Tiny brown beetle
JDN2010-29-4
Coleoptera
7/20/10
Wreck Pond
Tiny brown beetle
JDN2010-31-1
Hymenoptera
7/20/10
Wreck Pond
Spiny black ants
JDN2010-31-2
Hymenoptera
7/20/10
Wreck Pond
Spiny black ants
JDN2010-31-3
Hymenoptera
7/20/10
Wreck Pond
Spiny black ants
JDN2010-31-4
Hymenoptera
7/20/10
Wreck Pond
Spiny black ants
JDN2010-32
Coleoptera
7/20/10
Wreck Pond
smallish brown round beetle
JDN2010-33
Diptera
7/20/10
Wreck Pond
zebra wing fly
JDN2010-34
Coleoptera
7/21/10
Wreck Pond
FB2010-48-1
Hemiptera
7/22/10
FB2010-48-2
Hemiptera
7/22/10
FB2010-51
Hymenoptera
7/22/10
FB2010-52
Hymenoptera
7/22/10
FB2010-53
Hymenoptera
7/22/10
FB2010-54-1
Hemiptera
7/22/10
7/23/2010
FB2010-54-1 dup pcr
Hemiptera
FB2010-54-2
Hemiptera
FB2010-54-2 dup pcr
Hemiptera
FB2010-54-3
Hemiptera
FB2010-54-3 dup pcr
Hemiptera
FB2010-54-4
Hemiptera
FB2010-54-4 dup pcr
Hemiptera
7/23/2010
7/23/2010
7/23/2010
7/23/2010
7/23/2010
7/23/2010
7/23/2010
FB2010-56
Hymenoptera
7/23/2010
FB2010-58
Diptera
FB2010-7
Hemiptera
7/23/2010
very weak
+
+
+
+
+
medium black beetle
JDN2010-35-1
Diptera
7/21/10
Wreck Pond
+
zebra wing fly
JDN2010-35-2
Diptera
7/21/10
Wreck Pond
+
zebra wing fly
JDN2010-35-3
Diptera
7/21/10
Wreck Pond
JDN2010-35-4
Diptera
7/21/10
Wreck Pond
JDN2010-36-1
Diptera
7/21/10
Wreck Pond
longtail zebra wing fly
JDN2010-36-2
Diptera
7/21/10
Wreck Pond
longtail zebra wing fly
JDN2010-37
Hemiptera
7/21/10
Wreck Pond
Graphocephala versuta
JDN2010-38
Coleoptera
7/21/10
Wreck Pond
+
JDN2010-39
Coleoptera
7/21/10
Wreck Pond
+
JDN2010-40-1
Hemiptera
7/21/10
Wreck Pond
zebra face bug
JDN2010-40-2
Hemiptera
7/21/10
Wreck Pond
JDN2010-41-1
Coleoptera
7/21/10
Wreck Pond
JDN2010-41-2
Coleoptera
7/21/10
Wreck Pond
JDN2010-41-3
Coleoptera
7/21/10
Wreck Pond
zebra face bug
translucent wing, tiger striped
beetle
translucent wing, tiger striped
beetle
translucent wing, tiger striped
beetle
JDN2010-42
Hemiptera
7/21/10
JDN2010-47
Coleoptera
7/21/10
JDN2010-48
Hymenoptera
7/21/10
JDN2010-51
Coleoptera
7/21/10
JDN2010-55
Hymenoptera
7/21/10
JDN2010-61-1
Hymenoptera
7/22/10
JDN2010-61-2
Hymenoptera
7/22/10
JDN2010-61-3
Hymenoptera
7/22/10
JDN2010-61-4
Hymenoptera
7/22/10
JDN2010-61-5
Hymenoptera
7/22/10
JDN2010-61-6
Hymenoptera
7/22/10
JDN2010-62
Hymenoptera
7/22/10
JDN2010-63
Hymenoptera
7/22/10
JDN2010-66
Hymenoptera
7/22/10
JDN2010-67
Diptera
7/22/10
Wreck Pond
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
zebra wing fly
+
zebra wing fly
+
calico-checker tail bug
+
Harmonia axyridis
+
Dark green/blue wasp
medium black beetle (same as
morpho 34)
+
+
+
+
+
black/green small wasp
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
2 yellow spots on shoulder
black hornet, 2 yellow V's on face,
1 yellow dot on shoulder
black hornet, 2 white V's on face, 2
white dots on shoulder
small, dark green/black hornet,
long stinger
zebra wing fly
JDN2010-70-1
Coleoptera
7/22/10
JDN2010-70-10
Coleoptera
7/22/10
JDN2010-70-11
Coleoptera
7/22/10
JDN2010-70-2
Coleoptera
7/22/10
JDN2010-70-3
Coleoptera
7/22/10
JDN2010-70-4
Coleoptera
7/22/10
JDN2010-70-5
Coleoptera
7/22/10
JDN2010-70-6
Coleoptera
7/22/10
JDN2010-70-7
Coleoptera
7/22/10
JDN2010-70-8
Coleoptera
7/22/10
JDN2010-70-9
Coleoptera
7/22/10
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Manasquan
Cove
Small silver beetle
+
Small silver beetle
+
Small silver beetle
+
Small silver beetle
+
Small silver beetle
+
Small silver beetle
+
Small silver beetle
+
Small silver beetle
Small silver beetle
Small silver beetle
Small silver beetle
Figure 3. Proportion of PCR positives as a function of the total number of insects tested (n=96).
Figure 4. Proportion of PCR positives represented by each insect order.
Figure 5. Insect genomic DNA gels
Figure 6. PCR gel results.
Conclusion
According to previous data the expected amount of insects infected with Wolbachia is
around 20% and that about 60% of individuals in a species would be infected. We hypothesized
that the amount of our infected samples would be around the same percentage. After collecting
samples and running pcr test we found that out of the 96 individual samples tested that
approximately 30% were infected with Wolbachia. Out of the morpho species that were infected,
up to 86% of individuals were infected. These percentages were higher then expected percent,
however we were not able to conduct PCR testing all individual samples collected. In order to
test this hypothesize farther we would need to collect and test a larger number of samples and
test more replicates per species. . In addition we also asked if the location affected the amount of
infections. We saw that certain areas did have more infections then others but we would need to
do more testing to further characterize local ecological effects.
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